Scalable intracellular delivery via microfluidic vortex shedding enhances the function of chimeric antigen receptor T-cells

Abstract Adoptive chimeric antigen receptor T-cell (CAR-T) therapy is transformative and approved for hematologic malignancies. It is also being developed for the treatment of solid tumors, autoimmune disorders, heart disease, and aging. Despite unprecedented clinical outcomes, CAR-T and other engineered cell therapies face a variety of manufacturing and safety challenges. Traditional methods, such as lentivirus transduction and electroporation, result in random integration or cause significant cellular damage, which can limit the safety and efficacy of engineered cell therapies. We present hydroporation as a gentle and effective alternative for intracellular delivery. Hydroporation resulted in 1.7- to 2-fold higher CAR-T yields compared to electroporation with superior cell viability and recovery. Hydroporated cells exhibited rapid proliferation, robust target cell lysis, and increased pro-inflammatory and regulatory cytokine secretion in addition to improved CAR-T yield by day 5 post-transfection. We demonstrate that scaled-up hydroporation can process 5 x 108 cells in less than 10 s, showcasing the platform as a viable solution for high-yield CAR-T manufacturing with the potential for improved therapeutic outcomes.


INTRODUCTION
Adoptive chimeric antigen receptor (CAR) T-cell therapy requires the ex vivo modification of a patient's T-cells to express an engineered surface receptor that recognizes a unique tumor antigen via a single-chain variable fragment (scFv).Typically, this transmembrane receptor is linked to co-stimulatory intracellular signaling domains like CD3ζ paired with either 4-1BB or CD28 1,2 .After the stable modification has been conferred, CAR-Ts are expanded then re-infused into the patient where they specifically target tumor cells for lysis.Recent clinical studies demonstrated the efficacy of anti-CD19 CAR-T therapy with complete remission being reported in patients with acute or chronic lymphoblastic leukemias and B-cell lymphomas 1,3,4 .This led the U.S. Food and Drug Administration (FDA) to approve multiple CD19-directed CAR-T therapies for B-cell malignancies 5,6 .Despite the curative potential of these therapies, there are still major safety concerns regarding the toxicity associated with the powerful immune effector response induced by robust CAR-T activation, such as cytokine release syndrome (CRS) and immune effector cell-associated neurotoxicity syndrome (ICANS) 7,8 .These clinical results show the obvious potential of CAR-T therapy as treatment for hematologic malignancies, however, there are still several hurdles to overcome before its adoption as a first line treatment for many cancer types.Current manufacturing methods rely on lenti-or retroviral transduction for stable integration of the CAR transgene into the T-cell genome 9 .While this is a highly efficient approach, it lacks the precision editing that avoids insertional mutagenesis caused by random integration into the genome, potentially leading to oncogenic and mutagenic CAR-T products 10,11 .This raises serious safety concerns for clinical applications [12][13][14] .Furthermore, the strict regulations mandated for current Good Manufacturing Practice (cGMP) viral production laboratories make the process slow and expensive, which limits its feasibility for clinical-and commercial-scale manufacturing.
Precision genome editing overcomes these safety concerns by specifically integrating the transgene into a defined locus within the host genome.For example, clustered regularly interspaced short palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) CRISPR/Cas9 is a targeted nuclease that can induce a double stranded break (DSB) at a defined location 15,16 .By providing donor template DNA with homology arms complementary to the sequences that flank the cut site, the homology directed repair (HDR) pathway can then integrate the donor DNA sequence into that defined location 17,18 .In light of these advances, a series of seminal papers have been published which demonstrate the utility of CRISPR/Cas9 as a tool for adoptive T-cell engineering 19,16,15,20 .Generally, these approaches utilize electroporation ,or nucleofection, as a mechanism for intracellular delivery of gene editing payloads, where the cell membrane is permeabilized by exposing cells to one or more electric pulses of varying amplitude and duration.This creates pores in the plasma membrane by which the payload can enter the cell 21 .As demonstrated by Pal et al., 2024., use of a ribonucleoprotein (RNP) and adeno-associated viruses (AAV) CAR-T has shown significant promise in clinical trials, particularly in treatment of clear cell renal cell carcinoma (ccRCC) 22 .Pal et al., 2024., showed CD70-targeting CAR-T, could generate complete regression of ccRCC xenograft tumors in a mouse model prior to demonstrating 81.3% of patient outcomes resulted in disease control.CRISPR engineered CAR-T cells have also been used against CD19 for hematological malignancies.Stadtmauer et al., 2020, demonstrated in their phase 1 pilot, the safety and feasibility of using multiplex CRISPR-Cas9 T-cells 23 .Hu et al., 2021, also showed highly efficient, high-fidelity gene editing with universal CAR-T cells, with complete remission rate at 83.3% on day 28 24 .
Using electric pulses to permeabilize the plasma membrane remains one of the most common intracellular delivery methods to date.Both the Neon Transfection System (electroporation) and the Lonza 4D-Nucleofector (nucleofection) have been adopted as industry standards for intracellular delivery, particularly with hard-to-transfect cell types such as primary human T-cells, since they have unique buffers and pulsing protocols that are optimized for specific cell types 25,26 .While electroporation and nucleofection are efficient intracellular delivery mechanisms, there are severe drawbacks to this approach due to the significant damage caused by the electrical pulse 27 .Firstly, intracellular membrane-bound organelles are targeted by these pulses in addition to the plasma membrane.This results in the leakage of destructive enzymes from lysosomes, pro-apoptotic factors from mitochondria and various cytoplasmic components out of the cell 28,29 .Furthermore, electroporation has been shown to potentially cause irreversible genomic DNA damage as well as produce reactive oxygen species (ROS) which causes further damage to DNA, proteins and lipids within the cell [30][31][32][33] .Together, these effects put immense strain on electroporated cells, which can cause a high degree of cell death.
Herein, we describe hydroporation as an alternative intracellular delivery mechanism amenable to CAR-T generation for research and clinical applications.Hydroporation employs microfluidic vortex shedding (µVS), a hydrodynamic phenomenon whereby oscillating fluid forces gently permeabilize the cell membrane, allowing delivery of gene editing payloads such as Cas9 RNP.
Hydroporation relies on posts spaced approximately twice the typical cell diameter resulting in cell-size independent delivery and flexible throughput (ie. 10 4 ~10 8 activated T-cells mm -1 flow cell width) in a manner that is gentler than electroporation.Additionally, hydroporation utilizes 10~50x less reagent (i.e., mRNA, RNP) per cell than other nascent delivery platforms [34][35][36][37] .These fundamental characteristics of hydroporation lend themselves to cell therapy manufacturing where starting material between donors and patients can vary significantly, perturbation adversely affects cell function and a significant number of cells need to be processed 38,39 .We have previously demonstrated the utility of hydroporation as a mechanism for intracellular delivery of mRNA and Cas9 RNP targeting the TRAC locus in primary T-cells 40,41 .Here, we target TRAC disruption, through RNP delivery via hydroporation, with an anti-CD19 CAR AAV mediated homology directed repair template (HDRT) to achieve precision editing of CAR-Ts (Figure 1A) 16,42 .
In comparing the performance of hydroporation to nucleofection and electroporation, in generating CAR-Ts, we evaluated the transfection efficiency, viability, proliferation and total CAR-T yield.Hydroporation resulted in 1.7x to 2x greater yield of CAR-Ts on average by day 5 post-transfection compared to electroporation and nucleofection, respectively.To further demonstrate the versatility of hydroporation, we designed a high throughput microfluidic chip capable of processing up to 5 x 10 8 activated T-cells in less than 10 seconds, showing that hydroporation is scalable and capable of processing a wide range of cell numbers with similar yields in recovery, viability and editing efficiency.
We also sought to characterize the T-cell subset and cytokine secretion profile for CAR-Ts.Hydroporated CAR-Ts (Hy-CAR-Ts) and nucleofected CAR-Ts (Nuc-CAR-Ts) displayed nearly identical CD4+/CD8+ ratios as well as memory phenotype.Both Hy-CAR-Ts and Nuc-CAR-Ts demonstrated specific activation and pro-inflammatory cytokine secretion upon target cell engagement.
Finally, in order to evaluate the potency and clinical utility of Hy-CAR-Ts, we assessed their functionality using in vitro and in vivo models that mimic recent successful clinical trials utilizing AAV-mediated CAR-Ts, and compared them to Nuc-CAR-Ts 22 .We observed similar killing potential for Hy-CAR-Ts, though Hy-CAR-Ts showed lower levels of activation induced cell death (AICD) and improved motility in in vitro single cell serial killing assays.We observed equivalency between Hy-CAR-Ts and Nuc-CAR-Ts in an aggressive in vivo mouse xenograft model of B-cell acute leukemia.. Thus, hydroporation offers a means of generating high yields of precisely genome-edited CAR+ T-cells, reducing manufacturing costs and time needed to generate engineered cell therapies like CAR-T.

RESULTS
We have previously demonstrated the utility of hydroporation (i.e.µVS) as a mechanism for intracellular delivery of both mRNA and Cas9 RNP to primary human T-cells 40,41 .Importantly, previous studies indicated minimal dysregulation of the native T-cell state along with rapid cell recovery and proliferation following hydroporation.In order to evaluate the potential of hydroporation as a tool for adoptive immunotherapy manufacturing we compared hydroporation with nucleofection and electroporation in the generation of CAR-Ts.Using a previously validated CRISPR knock-in system, we utilized Cas9 RNP targeting the first exon of TRAC and an AAV vector encoding a self-cleaving P2A peptide, upstream from the homology directed repair template for TRAC, consisting of a CD19-specific 1928z CAR and EGFRt reporter 16 .Hydroporation, nucleofection or electroporation of activated primary human CD3+ T-cells took place 2 days after being isolated from frozen peripheral blood mononuclear cells (PBMCs).Cells were cultured for 9 days following RNP and AAV delivery and were evaluated for viability, proliferation, knockout (KO) efficiency, knock-in (KI) efficiency and CAR-T yield (Figure 1).
The fluid dynamic conditions created in the post array region of the microfluidic chip gently and efficiently permeabilize the plasma membrane to promote external material uptake (Figure 1B).Compared to electroporation and nucleofection, this method of membrane poration is less detrimental to cell health, which is reflected in the improved recovery and viability in hydroporated cells (Figure 1C and 1E).Hydroporated cells reach >90% viability within 2 days following transfection, while nucleofected and electroporated cells reach 90% viability after 5 days post-transfection.In terms of cell recovery, hydroporation and electroporation performed similarly, with 60-70% live cell recovery 2 hours after RNP delivery.Nucleofection showed the lowest recovery with <40% live cell recovery.
Both nucleofection and electroporation resulted in highly efficient KI rates (~80%), while hydroporation transgene insertion was lower, at ~40% (Figure 1G).However, in the 5 days following payload delivery hydroporated cells divided more rapidly, resulting in 3.2x and 3.6x more live cells than nucleofection or electroporation on average, respectively (Figure 1D and F).When taken with the corresponding recoveries and KI efficiencies, hydroporation yielded 1.7x and 2.0x more CAR-Ts, on average, than electroporation and nucleofection (Figure 1H).
Unlike static electric pulsing techniques, such as nucleofection or electroporation, which use cuvettes or pipette tips equipped with electrodes, hydroporation employs a stable flow delivery model which lends itself to be easily scaled up or down depending on the desired transfection volume (Figure 2A).Here we demonstrated the versatility of hydroporation by designing arrayed versions of our proprietary flow cell in order to accommodate greater numbers of cells.For small scale transfections (<10 8 cells) we used chips containing either 1 flow cell or 4 sub-flow cells which process cells simultaneously, known as Research Use Only (RUO) chips.For large scale transfections aimed at manufacturing a clinically relevant dose of CAR-Ts, we employed microfluidic chips which contained 40 arrayed flow cells, known as Cell Therapy (CT) chips.
When different numbers of cells are processed, there is a peak of around 40% CAR+ T-cells at 10 7 T-cells with the 4x flow cell RUO chip (Figure 2B), though there is no significant difference when processing higher cell numbers (10 9

cells showed around 25% CAR+ T-cell efficiency).
There is a trend where the greater the number of processed cells, the higher the % of TRAC KO (Figure 2C), particularly as we trend up to 5 × 10 6 cells.Cells at this density also show around 70% viability (Figure 2D), similar to that observed with electroporated and nucleofected cells.This is also observed in cell recovery, in which hydroporation recovers a higher number of transfected cells compared to both nucleofection and electroporation (Figure 2E).For hydroporation, it appears that higher cell numbers equates to greater recovery and transfection efficiency.Total sample processing times, for both the RUO and CT chips, range from milliseconds to seconds.For example, the RUO chip processes a typical sample of 5 x 10 6 activated T-cells (i.e., 100 µL at 5 x 10 7 cells mL -1 ) in less than 1 second, while the CT chip can handle 5 x 10 8 activated T-cells (i.e., 5 mL at 10 8 cells mL -1 ) in under 10 seconds.
To further understand the impact of the transfection method on T-cell phenotype, the proportions of naive and memory cell subsets in CD4 + and CD8 + T-cells were determined by flow cytometry for both Hy-CAR-Ts and Nuc-CAR-Ts.We did not observe any significant differences in the frequencies of these subpopulations in CD4 + or CD8 + T-cells from either donor between Hy-CAR-Ts and Nuc-CAR-Ts (Figure 3A).
To assess the function of Hy-CAR-Ts, we first characterized the secreted cytokine profile of the cells upon engagement with target NALM6, a CD19-expressing B-cell leukemia cell line, using nELISA.For this study, we co-cultured Hy-CAR-Ts, Nuc-CAR-Ts or wild type (WT) T-cells with NALM6 target cells overnight at a 1:1 ratio.The next day, supernatants were collected for protein quantification.For both donors tested, Hy-CAR-Ts and Nuc-CAR-Ts secreted high levels of pro-inflammatory cytokines, such as IL-2, IL-17A, and IFNɣ along with chemokines, such as IL-8, CCL1, CCL5, and the cytotoxicity effector molecule granzyme B, compared to WT samples (Figure 3B).These cytokine and chemokine expression patterns indicate robust activation via the anti-CD19 CAR.
We also compared the cytokine secretome of Hy-CAR-Ts directly with Nuc-CAR-Ts.While the number and magnitude of significantly different cytokines was relatively small, Hy-CAR-Ts displayed higher levels of cytokine secretion in both donors analyzed (Figure 3C).Interestingly, between the donors involved in this study, there was a common upregulated anti-inflammatory cytokine in activated Hy-CAR-Ts compared with Nuc-CAR-Ts -IL-10.Moreover, the pro-inflammatory chemokine IL-8 and the IL-10 family cytokine IL-22 were also upregulated in Hy-CAR-Ts compared with Nuc-CAR-Ts in both donors.
T-cell populations are highly variable, and cell phenotypes change after interaction with target cells.Therefore, we undertook a single-cell approach, Time-lapse Imaging Microscopy In Nanowell Grids (TIMING™) 43,44 , to compare the dynamics and functions of Hy-CAR-Ts with Nuc-CAR-Ts from each of five different donors (Figure 4).Cells are constrained within nanowells, allowing identification and tracking of individual cells over time before, during, and after interactions with other cells (Figure 4A).TIMING revealed differences between Hy-CAR-T and Nuc-CAR-T that cannot be detected in bulk assays 44 .We observed intrinsic donor variability in CAR-T target seeking and contact dynamics, but no significant differences in these parameters between CAR-T prepared by hydroporation and nucleofection.For example, the time required for CAR-T cells to form a stable synapse with target cells ("tSeek", not shown), the percent of effectors that formed at least one synapse, and individual synapse duration ("tContact") with target cells varied among donors, but there were no differences between the matched Hy-CAR-T and Nuc-CAR-T prepared from the same donor (Figure 4B).During target engagement (synapse), Hy-CAR-Ts derived from all five donors maintained significantly more motility than Nuc-CAR-T cells derived from the same donor (Figure 4C), consistent with prior data linking single-cell motility with resistance to exhaustion 45,46 .
Notably, the difference in CAR-T cell motility was not observed in the absence of target cells or prior to synapse formation.We observed no significant differences between Hy-CAR-T and Nuc-CAR-T cells in time required to kill targets, though there was substantial donor variability (Figure 4D).
A major advantage of TIMING is that by constraining cells in nanowells, the fate of effectors after target killing is quantifiable 47 .Consistent with lower motility during synapse, we found that Nuc-CAR-T were more sensitive to AICD than Hy-CAR-T cells from the same donor (Figure 4E).AICD occurred more frequently in Nuc-CAR-T irrespective of the number of targets engaged, but AICD in Nuc-CAR-T tended to increase with an increased number of targets engaged.The difference in AICD was also significant when survival times were compared by Kaplan-Meier analysis of three pooled donors (Figure 4F).Together single-cell results imply that CAR-T prepared by hydroporation are likely to resist exhaustion in vivo.
Recent literature suggests a strong correlation between CAR-T proliferation, in vivo persistence and improved patient outcomes 48 .Encouraged by the above results, we next sought to compare the cytotoxicity of Hy-CAR-Ts with Nuc-CAR-Ts using an in vitro co-culture assay with CD19-expressing NALM6 cells (Figure 5A).To enrich the CAR+ population to reduce noise in these studies, both Hy-CAR-Ts and Nuc-CAR-Ts were subject to TCR (T-cell receptor) depletion.
Hy-CAR-Ts demonstrated similar target cell lysis at all E:T ratios when compared with Nuc-CAR-Ts in this bulk killing assay (Figure 5B).In addition to the in vitro co-culture assay, an in vivo 'stress test', in which the CAR-T dose is lowered to reveal the functional limits of different CAR-T populations, was performed as previously reported 42 .A total of 10 5 or 4 × 10 5 CAR + T-cells, generated using either hydroporation or nucleofection, were injected intravenously into NOD.Cg-Prkdc scid Il2rg tm1Wjl /SzJ (NSG) mice engrafted with 5 × 10 5 NALM6 cells 4 days prior.An assessment on day 100 indicated the low dose of Hy-CAR-Ts had better survivability for donor 1, with a similar survival for donor 2 (Figure 5D).Tumor burden was evaluated over time by bioluminescence imaging (BLI) indicating a similar tumor burden for both transfection methods, at either CAR-T dose (Figure 5C).Taken together, these results demonstrate that Hy-CAR-Ts have similar therapeutic potency to their electroporation counterpart, which is typical for this preclinical model 42 .

DISCUSSION
The delivery of gene editing payloads to primary human T-cells continues to be an obstacle in the development of robust manufacturing methods for adoptive immunotherapy.Lenti-and retro-viral gene modification have severe limitations, including cost, safety, and scalability.Furthermore, gene delivery platforms that utilize electroporation or nucleofection have demonstrated a significant negative impact on cell health and function, which limits their potential for clinical applications.Recent studies that use Cas9-RNP show increasing utility in the specific integration of CARs or TCRs into the TRAC locus 49,50 .Other intracellular delivery methods exist, such as peptide-enabled RNP delivery for CRISPR (PERC) 42 , microneedle injection 34 , and cell squeezing 36 .However, these methods suffer from drawbacks like low throughput, high reagent consumption, and user safety concerns 51 .
These challenges motivated us to investigate microfluidic vortex shedding as a mechanism for cell membrane permeabilization and subsequent intracellular delivery of gene editing payloads.The advantages of the microfluidic transfection platform are evident in its gentle nature, ease of use, scalability, and reduced cost.Prior studies of hydroporation using mRNA 40 and Cas9 RNP 41 showed the potential for this platform as a tool for T-cell engineering, though the advantages were not fully realized until we achieved KI of an anti-CD19 CAR into the TRAC locus.When compared to nucleofection, not only did hydroporation yield a significantly greater number of CAR-Ts, but those CAR-Ts exhibited equivalent or better cytotoxicity in the presence of target cells.
To better understand the dynamics of this superior in vitro killing potential, we analyzed the T-cell phenotype and cytokine production.There was no difference in the composition of the naive/memory T-cell subsets between Hy-CAR-Ts and Nuc-CAR-Ts.Of the 187 cytokines tested, 3 showed significant difference between activated Hy-CAR-Ts and Nuc-CAR-Ts during killing for both donors: IL-8, IL-10, and IL-22.
IL-8 is a pro-inflammatory chemokine that attracts and activates neutrophils 52 .IL-8 production by tumor infiltrating activated CAR-T can thus increase neutrophil tumor infiltration.IL-8 is also secreted by several types of tumors, prompting researchers to generate CAR-T expressing the IL-8 receptors CXCR1 and CXCR2 to enhance CAR-T infiltration in solid tumors 53 .IL-22 is a pleiotropic cytokine, with reported pro-inflammatory and anti-inflammatory effects depending on the context 54 .In colorectal cancer, IL-22-producing infiltrating CD4+ and CD8+ T-cells were correlated with a better clinical outcome and increased infiltration of neutrophils, which in turn enhanced anti-tumor T-cell responses 55 .Moreover, CAR-Ts engineered to secrete IL-22 were found to be more cytotoxic towards head and neck squamous cell carcinoma 56 .
While the vast majority of cytokines produced upon T-cell activation are pro-inflammatory, IL-10 can act as a regulatory interleukin and suppress the immune response as a built-in negative feedback loop.IL-10 works by inhibiting the production of other pro-inflammatory cytokines, suppressing the antigen-presenting capacity of dendritic cells and macrophages, reducing the expression of major histocompatibility complex (MHC) class II and co-stimulatory molecules on antigen-presenting cells and promoting the differentiation of regulatory T-cells (Tregs) 57 .This suggests that even though Hy-CAR-Ts have an extremely robust pro-inflammatory cytokine response to CAR activation, there is still production of regulatory cytokines to limit excessive inflammation.
From an efficacy standpoint, the persistence of CAR-Ts in vivo is closely linked to positive clinical outcomes.CAR-T proliferation in vivo is fundamental to long term persistence.Studies indicate that electroporation limits the proliferative capacity of cells due to DNA damage as well as the generation of ROS caused by the electric pulse(s).As demonstrated by our results, hydroporated cells maintain a high proliferative capacity in vitro compared to nucleofected cells, which suggests improved persistence in vivo.CAR-T's persistence in vivo is far more nuanced than simple proliferative capacity.Recent literature shows that the tumor microenvironment can be very inhospitable to CAR-Ts, leading to AICD and exhaustion.Our results, though confined to a 6 hour window, show significantly lower levels of AICD with improved motility in CAR-Ts generated by hydroporation compared to nucleofection.This suggests that Hy-CAR-Ts are better equipped to robustly engage target cells without succumbing to AICD, which would potentially lead to longer persistence and a greater anti-tumor effect.

CONCLUSIONS & FUTURE WORK
We successfully used microfluidic vortex shedding to generate genome edited CD19-directed CAR-Ts with superior proliferative capacity, improved motility and reduced AICD when compared to nucleofected CAR-Ts.Hy-CAR-Ts were then shown to have (1) increased IL-10 production upon CAR engagement -a critical regulatory cytokine for CAR-Ts especially for solid tumors -all while (2) maintaining potency in vivo in a B-cell leukemia mouse xenograft model.We also demonstrated previously reported hydropore designs that maintain similar performance when scaled up and out to process 5 x 10 8 activated T-cells in less than 10 seconds -a key criterion for cell therapy manufacturing.
Cumulatively, these results indicate that hydroporation can be incorporated into traditional research and cell therapy manufacturing processes due to: (1) flexible throughput; (2) cell-size independent delivery; (3) gentle processing; (4) improved yields due to higher proliferation; (5)  reduced reagent consumption all (6) without loss of CAR-T function both in vitro and in vivo.
There are, however, still issues related to use of AAV for successful CAR KI due to size limitations.With this in mind, the hydroporation platform can be further developed to address these potential problems.Our future efforts are focused on multiplex editing and incorporating various forms of DNA to our KI strategies to exceed the size limitations of AAV.We see hydroporation being applied to other therapeutic, or difficult to transfect cell types, and we have already demonstrated the utility of hydroporation for regulatory CAR-Ts and natural killer cell KIs.

Chip fabrication & operation
Chips were fabricated using previously reported designs 40 and mask-based 3d printing (3d printed microtec, Bethesda, MD).Briefly, chips and chip features (i.e., inlet filter, posts, channel thickness, inlet and outlet channels) were manufactured by (1) generating a digital rendering of the microfluidic chip using Onshape CAD software, (2) preparing a photomask for patterning GR-1 resin (pro3dure, audioprint® GR-1) and ( 3) performing a series of 3d printing, metallisation, bonding and inspection steps.Chips were operated with a hydroporation instrument using with compressed nitrogen at 140 psig using previously reported protocols 41 .

Isolation & culture of primary human T-cells
Primary human CD3+ T-cells were isolated by negative selection from cryopreserved PBMCs (STEMCELL Tech, CAT#70025) using the EasySep Human T-cell Isolation Kit (STEMCELL Tech, CAT#17951) per the manufacturer's recommendation.After isolation, T-cells were seeded at 10 6 cells mL -1 in complete culture media.Complete culture media consisted of X-VIVO 20 media (Lonza, CAT#04-44Q) supplemented with 5% human serum AB (Access Biologicals, CAT#535-HI) and sterilized through a 20 nm vacuum filter.T-cells were stimulated for 48 hours with bead conjugated CD3/CD28 Dynabeads at a 1:3 bead-to-cell ratio for in vitro studies and a 1:1 bead-to-cell ratio ahead of the in vivo study (Thermo Fisher, CAT#11132D).48 hours after activation, Dynabeads are removed and T-cells are processed by Hydroporation, nucleofection or electroporation.Throughout the in vitro experiments, T-cells were maintained at a cell density of 10 6 cells mL -1 and supplemented with 50 IU mL -1 rh IL-2 (Peprotech, CAT#200-02) every 2 days.Ahead of the in vivo study, cells were cultured in X-VIVO 15 media containing 5% human serum, 5 ng ml −1 IL-7 (Miltenyi, CAT#130-095-367) and 5 ng ml −1 IL-15 (Miltenyi, CAT#130-095-760).

RNP assembly
RNPs were produced by combining target-specific sgRNAs (Synthego) and recombinant Cas9 (Truecut V2, Thermo Fisher Scientific, CAT#A36499).Briefly, lyophilized sgRNAs were reconstituted in Nuclease-free molecular grade water (Invitrogen, CAT#AM9938) to a concentration of 100 µM and then aliquoted for storage at -20°C.On the day of transfection, sgRNAs were thawed and diluted to equal volume of Cas9 in nuclease free water, then mixed with Cas9 at a 1:1.5 Cas9 to sgRNA molar ratio.The RNPs were complexed by incubating the sgRNA:Cas9 mixture for 15 minutes at 37C, then moved to room temperature until used in the transfection (<1hr).200 μg mL -1 RNP was then aliquoted into a clean eppendorf before adding cells for transfection via hydroporation, nucleofection or electroporation.
Transfection with RNP 48 hours after dynabead activation, T-cells were pelleted, washed with PBS, and thoroughly resuspended in OptiMEM (Gibco, CAT#31985062), P3 buffer with supplement (Lonza Bioscience, CAT#V4XP-3024), or R buffer (Thermo Fisher, CAT#MPK10025) at a cell density of 2 × 10 7 cells mL -1 .Before transfection, cells were added to eppendorfs containing RNPs for a total volume of 100 µL, and mixed thoroughly via pipetting.Nucleofection: For samples treated by nucleofection, following manufacturer guidelines, the 100 µL media consisting of cells, RNP in P3 buffer was added to 4D-Nucleofector cuvettes and treated using pulse code EH115.400 µL of complete culture media was added to each cuvette and then transferred to a tissue culture incubator for 15 minutes for cell recovery.After the recovery period, Nucleofected cells were seeded at 10 6 cells mL -1 in pre-warmed complete culture media and returned to the tissue culture incubator.For the in vivo study, cells were nucleofected in 20 µL reactions in the 96-well Lonza shuttle systems.Cell and payload concentrations were kept constant.Electroporation: For samples treated by electroporation, following manufacturer guidelines, electrolyte E2 buffer was added to the Neon Ⓡ tube, and allowed to come to room temperature .Using the 100 µL Neon Ⓡ electroporation tip, the 100 µL media consisting of cells, RNP in R buffer was transferred to the Neon Ⓡ tube containing E2 buffer and treated at 1600 V, 10 ms and 3 pulses.Treated cells were transferred to pre-warmed complete culture media and seeded at 10 6 cells mL -1 before being returned to the tissue culture incubator.Hydroporation: For in vitro experiments, hydroporation was performed at 160 psi and 28.3V (2.25 kV cm -1 ).For in vivo experiments, hydroporation was performed at 140 psi and 0V.Hydroporated samples were collected in 15 mL conical tubes at 10 6 cells mL -1 in pre-warmed complete culture media, then transferred to cultureware and kept in the tissue culture incubator.
In some cases, Nedisertib (M3814, MedChemExpress, CAT#HY-101570) was used at the manufacturer's recommended working concentration.Samples dosed with Nedisertib were centrifuged and resuspended in fresh, pre-warmed complete culture media 18-24 hours after dosing.All conditions were run in triplicate.

AAV dosing
For KI samples, AAV6 encoding a 1928z anti-CD19 CAR with an EGFRt reporter (Charles River) was added to cells, immediately after RNP delivery at an MOI of 20,000GC cell -1 , and incubated for 24 hours in serum free culture media at 37°C.After 24 hours, cells were centrifuged to remove AAV and seeded at 10 6 cells mL -1 in complete culture media.For CAR-Ts prepared for the in vivo mouse model, cells were centrifuged and resuspended in OptiMEM at 5 x 10 7 cells mL -1 .AAV was added at 20,000 MOI and cells were incubated for 1 hour at 37°C.Following the incubation, RNP was added to the cells at the concentration noted above, and processed via Hydroporation.For nucleofection samples, following the 1 hour AAV incubation cells were centrifuged.resuspended in P3 buffer, then RNP was added and the cells were subjected to nucleofection according to the above methods.Following RNP delivery, cells were collected in complete culture media containing 5% human serum.

Flow cytometry
Transfected cells at different times post-transfection were analyzed by flow cytometry to measure the cell concentration, viability, KI and KO efficiency.All reagents were used according to manufacturer's recommendations.To measure viability and cell concentration, cells in media were diluted 1:20 in PBS containing propidium iodide (Sigma-Aldrich, CATt# P4170) then run on the flow cytometer.To measure KI and KO efficiency, cells were pelleted, washed with PBS, and gently resuspended and incubated for 30 min at 4°C in the antibody staining cocktail.The staining cocktail was composed of anti-CD3 FITC (Invitrogen, clone UCHT1) and anti-EGFR APC (Bioloegend, clone AY13) monoclonal antibodies in FACS buffer.After incubation, cells were washed in FACS buffer, pelleted, and resuspended in FACS buffer containing SYTOX Blue viability stain (Thermo Fisher Scientific, CATt# S34857).Samples were then acquired using an Attune NxT flow cytometer (Thermo Fisher Scientific).Compensation was performed using single-stained controls prepared with AbC Total Antibody Compensation Bead Kit (Thermo Fisher Scientific, CAT#A10513).Flow cytometry standard files were exported and analyzed using FlowJo software v3.0 (FlowJo).A standard gating strategy was used to remove debris and aggregated cells.Dead cells were excluded based on SYTOX Blue viability staining.

TCR magnetic depletion
Two days before performing the cytotoxicity assay or infusing into mice, CAR-T populations were enriched by depleting TCRɑ/β+ cells using the Miltenyi human TCRɑ/β depletion kit (Miltenyi CAT#130-133-896).Prior to selection, cell density and viability were assessed using a Cellaca ® PLX system (Nexcelom Bioscience).Cells were then centrifuged, resuspended in MACS buffer (80 μL per 10 7 cells, PBS, 0.5M EDTA, 2% Human serum), and incubated with a biotin-conjugated anti-TCRɑ/β antibody (20 μL per 10 7 cells) for 10 minutes at 4°C.Cells were then washed, resuspended in MACS buffer (80 μL per 10 7 cells), and incubated with anti-biotin microbeads (20 μL per 10 7 cells, Miltenyi) for 15 minutes at 4°C.Labeled cells were loaded onto LS Miltenyi MACS columns and processed according to the manufacturer-provided protocol.Cell density in the flow-through from the column was assessed, and isolated cells were centrifuged and resuspended in complete T-cell medium for culture.

Cytotoxicity assay
The cytotoxicity of anti-CD19 CAR-T cells was determined by standard luciferase-based assay.In brief, a stable NALM6-Fluc/eGFP cell line served as target cells.The effector (E) and tumor target (T) cells were co-cultured in triplicates at the indicated E:T ratio (1:1 to 1:64) using white-walled 96-well flat clear-bottom plates with 5x10 4 target cells in a total volume of 100 μL per well in complete T-cell media.The control for maximum signal was NALM6 cells alone, and the control for minimum signal was NALM6 cells and Tween-20 (0.2%).Co-cultures were incubated for approximately 22 h.Then, 100 μL D-luciferin (GoldBio, 0.75 mg ml −1 ) was added to each well, and luminescent signal was measured using a GloMAX Explorer microplate reader (Promega).Cytotoxicity = 100% × (1 − (sample − minimum)/(maximum − minimum)).

nELISA cytokine Multiplex Assay
To measure secreted cytokine concentration, after an overnight co-culture of CAR-T cells with NALM6 cells (1:1 E:T), 50-100 µL of cell supernatant was collected in a 96 well plate and briefly centrifuged to remove cells and debris.Cleared supernatants were then frozen in a -80°C freezer prior to overnight shipping to Nomic Bio (Montreal, Canada) for subsequent analysis.
Upon arrival, supernatants from treated cells were thawed for nELISA-based secretome analysis using standard protocols, as described previously 59 .Briefly, the nELISA pre-assembles antibody pairs on spectrally encoded microparticles, resulting in spatial separation between non-cognate antibodies, preventing the rise of reagent-driven cross-reactivity, and enabling multiplexing of hundreds of ELISAs in parallel.Protein concentrations on microparticles were read out by high-throughput flow cytometry (Bio-Rad ZE5 cell analyzer) and decoded using Nomic's proprietary software.Standard curves for all targets were generated to derive pg mL -1 values from cytometry fluorescence units.The nELISA MaxPlex panel was used to quantify 187 analytes in each sample.
nELISA Cytokine Multiplex Assay Data Analysis Data analysis was conducted in R (version 4.2.2) with the following packages: tidyverse (v.), data.table(v.), EnhancedVolcano (v.), heatmaply (v.), and reticulate (v.), with associated dependencies within RStudio (v.2022.12).The data analysis workflow was as follows.Raw values from the nELISA were read into R after reformatting in Microsoft Excel to remove extraneous rows to produce a .csvfile containing relevant data rows and the concentration (in picograms per mL) or raw nELISA signal.These were then statistically analyzed for comparisons of interest using one-way ANOVA.The raw signal values were used to calculate the statistical significance, as these values are more dynamic and more closely represent variation in the signal while fold change was calculated based on the transformed concentration values to incorporate data relevant to the underlying biology.The resulting log2fc and p-values were plotted on a volcano plot using the EnhancedVolcano package.These data were also used to generate heatmaps using the heatmaply package by selecting proteins at each condition meeting the following thresholds: ≥|log2fc| and p-value < 0.01.This more stringent p-value was selected to account for multiple comparisons within the data and to reduce the potential for incorrectly calling changes in these proteins.These were then matched to corresponding GO terms, retrieved from Uniprot and DICEDB.Heatmaps were generated using the heatmaply package.These changes were then summarized into bar plots using ggplot by calculating the numbers of proteins meeting these parameters: "up" encompassed proteins with a p-value < 0.01 and a log2fc > 2, "down" included proteins with a p-value < 0.01 and a log2fc <-2, and "same" includes all remaining proteins that fail to meet these criteria.

Animal Study
After 2 days of Dynabead activation, T-cells were edited by nucleofection or hydroporation as described above for the KI of a 1928z anti-CD19 CAR with an EGFRt reporter at the TRAC locus (M3814 was used for this experiment).TCR depletion was performed 5 days after editing and flow cytometry was conducted subsequently to estimate the CAR%.NOD.Cg-Prkdc scid Il2rg tm1Wjl /SzJ (NSG) mice were handled ethically and in accordance with the protocol AN182757-01G approved by the University of California, San Francisco (UCSF) Institutional Animal Care and Use Committee.Before and during the experiment, mice were maintained on Clavamox antibiotic.A total of 5 × 10 5 NALM6 cells were injected into the tail vein of mice that were between 8 and 12 weeks old.After the first BLI measurement (after NALM6 injection, the day before T-cell injection), mice were assigned to each T-cell condition so as to maintain a similar average mass and tumor burden across conditions.Four days after NALM6 injection, 10 5 or 4 × 10 5 CAR+ T-cells or an equivalent total number of nucleofected TCR KO T-cells were injected into the tail vein.Mouse health and survival were monitored over time.BLI was performed one or two times per week using a Xenogen in vivo imaging system.At each imaging session, mice were injected intraperitoneally with luciferin (3 mg luciferin per 0.2 ml DPBS) and anesthetized with isoflurane (Medline Industries, CAT#66794-0017-10).The default imaging exposure was 1 min, and shorter exposures were used for images that had a saturating signal at 1 min.Luminescence was quantified using Living Image software (PerkinElmer).Reported BLI values are an average from imaging each mouse on its front and on its back.Mice were euthanized per the approved protocol in the event that they reached end points such as loss of mobility or other signs of morbidity.

Figure 2 .
Figure 2. Scaled up version of the hydroporation chip, shows similar TRAC-1 KO, viability and recovery when compared to electroporation and nucleofection even when processing 1 billion activated T-cells.A) CAD images of the small volume RUO chip (1x flow cell), standard volume RUO chip (4x Flow cell) and large volume CT chip (40x Flow cell), with an enlarged image showing the post design per flow cell.B) Physical representation of CT chip (40x flow cells, left) to RUO chip (1-4x flow cells, right), bar 5 mm.C) Percentage of CAR+ T-cells on day 5 after transfection of10 6 -5 × 10 8 total cells.D) TRAC KO% efficiency, against cell numbers, based on whether cells were hydroporated (Hydro; blue), electroporated (EP; yellow) or nucleofected (Nuc, Magenta).E) TRAC KO Cell viability 24 hours post-transfection.F) Cell recovery 2 hours post transfection.All data points involve n = 3 biological replicates except 40x FC data.

Figure 3 .
Figure 3. T-cell phenotype and cytokine profile show no significant differences between hydroporation and nucleofection.A) Relative proportions of naive/memory CD4 + and CD8 + T-cell subsets for CAR+ cells engineered via hydropore (Hy-CAR-T) or nucleofection (Nuc-CAR-T).Percentages of naive/memory cells for both CD4 + and CD8 + T-cells were calculated based on expression of CD45RA and CD62L.B) Heatmap of 40 proteins secreted by CAR-T cells that were significantly differentially expressed compared to WT cells in the presence of NALM6 target cells.C) Comparison of cytokine secretion patterns of CAR-T cells from donor 1 and donor 2 generated using either nucleofection or hydroporation upon activation with NALM6 target cells at a 1:1 ratio, as assessed by 187-plex nELISA.NS: not significant; FC: fold-change.

Figure 4 .
Figure 4. CAR-T cells manufactured with hydroporation have superior motility and resistance to AICD.A) Images of cells in TIMING nanowells at different E:T ratios at the beginning of TIMING assays (00:00 hh:mm).CAR-T effectors (green), NALM6 target cells (magenta), bars 20 µm.B) Violin plots show population dynamics of synapse duration (tSynapse) between Hy-CAR-T (blue) or Nuc-Car-T (magenta) and NALM6.Bottom line indicates percent of CAR-T observed to form a synapse (Syn) in each assay.C) CAR-T motility (µm/ min displacement) before or during synapse with target cells.Each point is the mean for each donor (D1-D5), shown as paired samples of Hy-CAR-T (blue) vs Nuc-CAR-T (magenta).*p<0.05 by pairwise Mann-Whitney tests.D) Time required for CAR-T cells to kill target cells (tDeath of target, min) after synapse formation.Bottom line shows percent of synapses resultingin a kill.tDeath distributions: *p<.05, **p<0.01,***p<0.001,****p<.0001by pairwise Mann-Whitney.Kill % difference: *p<.05 by Fisher's exact test on raw numbers (only D5 showed differential killing of Hy-CAR-T vs Nuc-CAR-T.E) Mean AICD of effectors after engagement with one or more target cells (x-axis).Each connected pair of points indicates an individual donor.**p<.05,**p<0.01 by 2-tailed t-tests.F) Survival curves of effectors after target contact (AICD) over 6-hour TIMING assay.Curves represent pooled data from three donors (D3-D5).****p<.0001by Kaplan-Meier analysis.

Figure 5 .
Figure 5. CAR-Ts modified through hydroporation showed similar yields of CAR+ cells as nucleofected T-cells, as well as similar levels of cytotoxicity, but improved functionality as shown in survival rates of mice at Day 100.A) CAR-T+% after TCR depletion for donors 1 and 2 transfected by hydroporation or nucleofection.B) Bulk cytotoxicity assay, based on treated T-cells at different ratios of effectors to target cells (CAR-T to NALM6).C) BLI values, for donors 1 and 2, on the last measurement day on which all mice were alive (Day 13) and which all CAR-T injected mice were alive (Day 32).P values are from two-tailedWelch's unpaired t-tests.D) Kaplan-Meier survival analysis of mice treated with/without hydroporated or nucleofected CAR+ T-cells, over 100 days.N = 8 mice per group D) BLI values over 100 days.